Download Somatic Cytokinesis and Pollen Maturation in Arabidopsis

The Plant Cell, Vol. 18, 3502–3518, December 2006, www.plantcell.org ª 2006 American Society of Plant Biologists
Somatic Cytokinesis and Pollen Maturation in Arabidopsis
Depend on TPLATE, Which Has Domains Similar
to Coat Proteins
W
Daniël Van Damme, Silvie Coutuer, Riet De Rycke, Francois-Yves Bouget,1 Dirk Inzé, and Danny Geelen2,3
Department of Plant Systems Biology, Flanders Interuniversity Institute for Biotechnologie, Ghent University,
B-9052 Gent, Belgium
TPLATE was previously identified as a potential cytokinesis protein targeted to the cell plate. Disruption of TPLATE in
Arabidopsis thaliana leads to the production of shriveled pollen unable to germinate. Vesicular compartmentalization of the
mature pollen is dramatically altered, and large callose deposits accumulate near the intine cell wall layer. Green fluorescent
protein (GFP)–tagged TPLATE expression under the control of the pollen promoter Lat52 complements the phenotype.
Downregulation of TPLATE in Arabidopsis seedlings and tobacco (Nicotiana tabacum) BY-2 suspension cells results in
crooked cell walls and cell plates that fail to insert into the mother wall. Besides accumulating at the cell plate, GFP-fused
TPLATE is temporally targeted to a narrow zone at the cell cortex where the cell plate connects to the mother wall. TPLATEGFP also localizes to subcellular structures that accumulate at the pollen tube exit site in germinating pollen. Ectopic
callose depositions observed in mutant pollen also occur in RNA interference plants, suggesting that TPLATE is implicated
in cell wall modification. TPLATE contains domains similar to adaptin and b-COP coat proteins. These data suggest that
TPLATE functions in vesicle-trafficking events required for site-specific cell wall modifications during pollen germination
and for anchoring of the cell plate to the mother wall at the correct cortical position.
INTRODUCTION
The cell wall of higher plant cells provides the mechanical
strength required to hold the structure of the entire plant body.
New walls are laid down after mitosis through the activity of a
cytoskeletal configuration known as the phragmoplast. The
immature wall or cell plate emerges first at the cell center from
deposits that travel along microtubules to the spindle midzone. In
the next step, the young disk-shaped plate expands outward
until it reaches the cellular boundaries and unites with the existing
mother wall (for review, see Jürgens, 2005a, 2005b). Positioning
of the new cell wall after mitosis is critical for the establishment of
the cellular organization of plant tissues and the overall morphology of the plant (Lloyd, 1995; Traas et al., 1995). Plants have
developed regulatory mechanisms to position and guide the new
cell plate. Two plant-specific microtubular arrays are involved in
the positioning and guidance of the new cell plate, the prepro-
1 Current address: Laboratoire Arago, Unité Mixte de Recherche 7628,
Centre National de la Recherche Scientifique, Université Pierre et Marie
Curie, B.P. 44, F-66651 Banyuls sur Mer cedex, France.
2 Current address: Department of Plant Production, Faculty of Bioscience and Bioengineering, Ghent University, Coupure Links 653, B-9000
Gent, Belgium.
3 To whom correspondence should be addressed. E-mail danny.
[email protected]; fax 32-9-264-62-25.
The author responsible for distribution of materials integral to the
findings presented in this article in accordance with the policy described
in the Instructions for Authors (www.plantcell.org) is: Danny Geelen
([email protected]).
W
Online version contains Web-only data.
www.plantcell.org/cgi/doi/10.1105/tpc.106.040923
phase band (PPB) and the phragmoplast itself. The PPB consists
of a ring of microtubules and actin filaments, encircles the
nucleus at prophase, and determines the future division zone
(Mineyuki and Gunning, 1990; Wick, 1991; Mineyuki, 1999). The
PPB is removed before chromosome segregation and cytokinesis and therefore cannot contribute directly to the cell plate guidance process. As the PPB microtubules degrade, cortical actin at
the corresponding former position of the PPB disappears, leaving behind a zone devoid of actin filaments (Cleary et al., 1992;
Sano et al., 2005). The role of actin in cytokinesis is not very clear.
Actin depolymerization slows mitotic progression and affects
phragmoplast initiation. However, it does not have a major
impact on cell plate positioning (Yoneda et al., 2004). Recently,
a novel marker was identified that implicates the plasma membrane as an additional structure that may be important for determining the division zone (Vanstraelen et al., 2006). The marker
is a green fluorescent protein (GFP)–tagged kinesin, KCA1, which
associates with the plasma membrane at the onset of mitosis. It is
excluded from a region corresponding to the size and position of
the PPB and the actin-depleted zone. By analogy, it was called
the KCA-depleted zone or KDZ. The KDZ persists throughout
mitosis until the new cell plate has expanded to reach the mother
wall and therefore marks the division zone. It remains to be
investigated whether KCA1 plays a direct role in the positioning
and guidance of the cell plate.
The phragmoplast is required to construct the new cell plate
and conducts guidance of the plate to the division zone. The double
array of parallel-oriented microtubules of the phragmoplast transport Golgi- or endosome-derived vesicles, containing cell plate–
building blocks, to the equatorial plane of the cell, where they
Cytokinesis and Pollen Depend on TPLATE
immediately fuse and give rise to a tubulovesicular network.
Callose deposition inside the lumen of the network may provide a
spreading force for the widening of the tubular network and
convert it to a fenestrated sheet (Samuels et al., 1995). Fusion of
the cell plate with the plasma membrane triggers a maturation
process that involves the replacement of callose by cellulose.
The mechanism and controlling elements required for the switch
from a callose- to a cellulose-containing cell plate are still unknown.
Mutants that are altered in cellulose biosynthesis often show
cytokinesis defects: cyt1 (Nickle and Meinke, 1998; Lukowitz et al.,
2001), prc1 (Fagard et al., 2000), kor1/rsw2 (Zuo et al., 2000;
Lane et al., 2001), kob1 (Pagant et al., 2002), and fk, hyd1, and
smt1/cph (Schrick et al., 2000). The cytokinesis defects described for cyt1, kob1, prc1, fk, hyd1, and smt/cph can be attributable to impeded wall formation that is manifested by disrupted
cell walls in microscopic sections. The kor1-2 mutant produces,
in addition to cell wall stubs, curved and misplaced cell walls,
despite the fact that the PPB is correctly positioned in these cells
(Zuo et al., 2000). korrigan (kor1-2) is an endo-1,4-b-glucanase,
indicating that factors that are not linked directly to PPB activity
are also required for the correct guidance of the cell plate.
Ultimately, cell plate expansion results in a fusion with the
existing wall. From a mechanistic viewpoint, the cell plate inserts
in the mother wall through a multitude of finger-like fusion tubes
that contact the parental plasma membrane in the zone of
adhesion (Samuels et al., 1995). Because the cell plate membrane and the plasma membrane are at first independent structures, their unification must rely on a membrane fusion process
that involves membrane fusion machinery different from the
KNOLLE/KEULE/SNAP33 SNARE complex that is required for
cell plate formation and expansion (Jürgens, 2005a). To date,
there is little information on the components that contribute to the
anchoring of the plate to the mother wall. A candidate protein is
ROOT/SHOOT/HYPOCOTYL-DEFECTIVE (RSH), a hydroxyl
Pro-rich glycoprotein (extensin) that accumulates at the site of
cell plate–cell wall contact. Disruption of the RSH protein is
embryo-lethal and causes misplaced cell plates, resulting in
irregular cell shape and size (Hall and Cannon, 2002).
Here, we report the functional analysis of TPLATE, a gene with
a role in cell plate anchorage. TPLATE-GFP (previously T22-GFP)
localizes to the midline of the expanding phragmoplast in dividing tobacco (Nicotiana tabacum) BY-2 cells (Van Damme et al.,
3503
2004a). During the final steps of cell plate expansion, GFP-fused
TPLATE accumulates at a defined zone of the plasma membrane
that corresponds to the division zone. TPLATE-GFP also accumulates at the pollen tube exit site during pollen germination, in
agreement with the male-sterility phenotype that is caused by a
T-DNA insertion in TPLATE. How cytokinesis and pollen development are connected is inferred from the similarity of TPLATE
and vesicle-associated proteins, suggesting a role in heterotypic
vesicle fusion.
RESULTS
TPLATE Is Similar to Coat Proteins
A GFP-based screen in BY-2 cells identified TPLATE (clone T22)
as a putative cell plate–targeted protein (Van Damme et al.,
2004a). To analyze the relevance of this gene to other species, the
occurrence of homologous sequences was determined (Figure
1). TPLATE is unique to plant species and occurs as a singleton
located on chromosome 3 in Arabidopsis thaliana. There are two
copies in rice (Oryza sativa var japonica), one on chromosome 11
(Os11g07470) and one on chromosome 2 (Os2g55010). Arabidopsis TPLATE and the rice homologs exhibit a genomic structure of seven exons and six introns (Figure 2). The Arabidopsis
TPLATE is ;70% identical to the rice homologs and 82%
identical to a predicted protein sequence from Lotus corniculatus
var japonicus (AP004906). A partial EST sequence (translated 160
amino acids) from Physcomitrella patens (BJ187435) is 57%
identical (79% similar), indicating that the protein is highly conserved from mosses to higher plants. The TPLATE open reading
frame predicts a protein with a molecular mass of 131 kD and
contains domains with similarity to an EF-hand motif (IPR002048)
and an adaptin_N domain (IPR002553), as identified by standard analysis (www.sanger.ac.uk/cgi-bin/pfam; Figure 2A). The
adaptin_N domain is present in the N-terminal part of the large
subunits of the AP-1, AP-2, AP-3, and AP-4 adaptor protein
complexes and in the b-COP, g1-COP, and g2-COP subunits of
the COPI protein complex (Boehm and Bonifacino, 2001). Adaptin protein complexes (AP) and coat protein complexes (COP) are
involved in clathrin-coated and non-clathrin-coated vesicle formation (McMahon and Mills, 2004). Besides an adaptin_N-like
domain, TPLATE carries a stretch of 14 amino acids highly
Figure 1. Alignment of the b-COP–Specific Element.
A stretch of 14 amino acids (the b-COP–specific element) is conserved in b-COP and TPLATE. Conserved amino acids in b-COP proteins derived from
different species are shown in boldface. Numbers represent the amino acid positions of the element within the proteins.
3504
The Plant Cell
Figure 2. Analysis of tplate Mutant Pollen.
Cytokinesis and Pollen Depend on TPLATE
conserved in b-coatomer proteins (b-COP). We called this motif
the b-COP–specific element (Figure 1). The composition of the
motif was determined as [P]L[T]G-[S]-SDP-x-Y-x-E[AY], with x
indicating any amino acid and amino acids in brackets partially
conserved in the b-COP protein family. The b-COP–specific
element is conserved in b-COP proteins across the different
genera (Figure 1). The occurrence of a b-COP element and a
domain similar to adaptin_N in TPLATE suggests that the TPLATE
protein is related to coat proteins.
A T-DNA Insertion in TPLATE Causes Complete
Male Sterility
TPLATE is interrupted at the third intron by a T-DNA insertion in
the Arabidopsis line SALK_003086 (Figure 2A). Sequences adjacent to the insertion site are amplified by PCR with the T-DNA
left border primer (LBaI) and primers of neighboring sequences
(T22-RP and T22-LP), suggesting that the insertion is an inverted
repeat. Determination of the T-DNA–bordering sequences confirmed the presence of two left borders and identified the exact
position of the insertion site (Figure 2A). Segregation analysis of
heterozygous plants resulted in 50% progeny showing resistance to kanamycin. All kanamycin-resistant plants (n ¼ 30) were
heterozygous for the T-DNA insertion. PCR on a population of
plants that were grown without selection also yielded the 1:1
segregation ratio (131 heterozygous and 151 wild-type plants; x2
test for a 1:1 ratio yields 1.42, and x2 95% interval value is 3.84).
Pollination of wild-type plants with pollen from the TPLATE
heterozygous mutant resulted solely in kanamycin-sensitive
plants (72 plants tested), indicating that the mutation cannot be
passed on by the male gametophyte and that the T-DNA insertion in TPLATE affects pollen development or germination. By
contrast, the insertion mutation has no discernible effect on the
development of the female gametophyte, as seed setting and
yield were similar to those of wild-type plants (data not shown).
tplate Mutant Pollen Shows Normal Karyokinesis and
Is Defective in a Late Developmental Stage
The development of the male gametophyte was analyzed using
light and fluorescence microscopy. Pollen produced by a heter-
3505
ozygous tplate mutant plant is shown in Figure 2. Dehiscing
anthers were stained with Alexander’s stain to discriminate
between live and dead pollen. The cytoplasm of viable pollen
becomes brightly red, whereas dead pollen is not stained by the
dye (Alexander, 1969). Two types of pollen grains were observed:
wild-type pollen that is oval-shaped, and shriveled, irregularly
shaped mutant pollen (Figures 2M and 2N). The ratio between
irregular and normal pollen was 1:1 (532 mutant and 526 wildtype grains; x2 test for a 1:1 ratio yields 0.03, and x2 95% interval
value is 3.84). The cytoplasm of the mutant pollen grains stained
red with the Alexander stain, indicating that the pollen is not dead
at this stage. Yet, some of the pollen had collapsed and showed a
thick layer of translucent deposit against the pollen intine wall
(Figure 2N).
To determine the developmental stage and nuclear composition of the pollen, dehiscing anthers were stained with 49,6diamidino-2-phenylindole (DAPI). Wild-type (Figure 2J) and
mutant (Figure 2K) pollen carried three nuclei, two male gamete
nuclei and one vegetative nucleus, indicating that nuclear divisions occurred during pollen development. Observation of earlier
stages in pollen development did not reveal morphological
differences and tetrads, as ring-vacuolated, bicellular, and early
trinucleate pollen appeared as in wild-type plants, suggesting
that the mutation affects later stages of pollen maturation (data
not shown). To assess the germination capacity of wild-type
versus mutant pollen grains, anthers were spread on germination
medium and observed after 8 and 16 h of incubation. In a control
experiment, 80% of wild-type pollen germinated. Pollen derived
from anthers harvested from mutant plants showed a reduction
of germination capacity of ;50%, and none of the shriveled
pollen germinated (Figure 2O).
Callose Accumulates Ectopically in Mutant Pollen Grains
The thick deposits at the cell periphery in mutant pollen
grains may have prevented germination by changing the structure of the intine wall layer. To determine the structure and
composition of the mutant pollen cell wall, sections of plasticembedded whole-mount anthers were stained with fluorescent
cell wall dyes. Propidium iodide (PI) treatment identified three
Figure 2. (continued).
(A) TPLATE gene structure with indication of domains, the T-DNA insertion position, and the position of the tobacco cDNA AFLP tag (BSTT43-4-330)
used for RNAi in BY-2 cells. Arrows indicate primers (Lba1, T22 LP, and T22 RP) used to check the T-DNA insertion. Sequence information obtained by
sequencing the fragments (T22 LP–Lba1 [underlined] and T22 RP–Lba1 [double underlined] primer pairs) identifies the T-DNA insertion within the third
intron and confirms the inverted repeat T-DNA structure.
(B) to (E) Confocal images of thin sections through anthers of a TPLATE heterozygous plant. Thin sections (5 mm) were stained with PI. Mutant pollen in
images (C) and (D) show inclusions not stained by PI. (E) shows a shriveled pollen grain.
(F) to (I) Epifluorescence images of thin sections, costained with calcofluor white (blue) and aniline blue (yellow), showing a wild-type pollen grain (F) and
callose depositions ([G] to [I]) inside the mutant pollen of images (B) to (E).
(J) and (K) Epifluorescence images of wild-type (J) and mutant (K) pollen stained with DAPI. Mutant pollen is trinucleate. gn, gamete nucleus; vn,
vegetative nucleus. Bars ¼ 20 mm.
(L) Scanning electron micrograph of mutant and wild-type pollen grains. Bar ¼ 10 mm.
(M) and (N) Bright-field microscope images showing an overview (M) and a close-up (N) of wild-type and mutant pollen grains visualized with
Alexander’s stain. Bars ¼ 100 mm in (M) and 20 mm in (N).
(O) Germinating pollen grains. Normal-looking pollen germinates (black arrowheads), in contrast with the mutant pollen (white arrowheads).
Bars ¼ 20 mm.
3506
The Plant Cell
types of pollen grains. Fifty percent of the pollen grains had a
normal ellipsoidal morphology and fluoresced homogenously
(Figure 2B). The other half of the pollen grains showed random
deposits of material that was not stained by PI (Figures 2C and
2D). In severely affected pollen with a shriveled appearance,
there was no PI staining (Figure 2E). This type of pollen occurred
more frequently in mature anthers and probably represents an
end stage in the maturation of the mutant pollen. The deposits
were also observed in the representative bright-field images
(Figures 2C and 2D). The deposits occurred mostly at the cell
periphery; therefore, the intine cell wall composition was analyzed using calcofluor white and aniline blue staining methods. A
mixture of aniline blue and calcofluor white distinctively labels
callose (yellow fluorescence) and cellulose (blue-green fluorescence), respectively (Figures 2F to 2I) (Jefferies and Belcher,
1974). The wild-type intine cell wall was predominantly composed of cellulose and stained brightly blue with calcofluor
(Figure 2F). The intine wall from shriveled pollen (Figures 2G to
2I) was fluorescent in the presence of calcofluor white, albeit
less pronounced than in wild-type grains. Hardly any aniline
blue fluorescence occurred in wild-type pollen (Figure 2F) because very little callose was present in these cells. By contrast,
brightly yellow fluorescent spots occurred in mutant pollen
(Figures 2G to 2I).
To analyze the deposits in more detail, pollen morphology was
determined by means of scanning and transmission electron
microscopy. The scanning images revealed that the exine layer of
mutant pollen is similarly structured as in wild-type pollen (notice
that the mutant pollen grain is approximately half the size of the
wild-type pollen grain; Figure 2L). Transmission electron microscopic analysis further confirmed that the structure and organization of the exine are normal. Figure 3A shows an overview of an
ultrathin section of a mutant and a wild-type pollen grain. Wildtype mature pollen is densely packed with vesicles, endoplasmic
reticulum, and Golgi (Figures 3B and 3E). At the periphery, a band
of Pb acetate–stained vesicles concentrates adjacent to the
intine layer. Closer to the center, there are more electron-dense
(dark) vesicles mixed with less numerous (electrolucent) vesicles
(Figure 3B). The peripheral vesicles are presumably involved in
secreting polysaccharide cell wall components upon germination and subsequent pollen tube growth (Heslop-Harrison, 1987).
Mature mutant pollen grains show an altered composition of the
pollen cytoplasm with small and large electron-dense bodies
(Figures 3B and 3H). The composition of the cytoplasm in not fully
matured mutant pollen with less severe morphological deformations shows an intermediate stage with undulations of the plasma
membrane (arrows in Figures 3C and 3G). Using a specific
antibody directed against callose, strong labeling was detected
specifically between the intine cellulose layer and the undulated
plasma membrane (Figure 3D), whereas hardly any signal was
detected in mature wild-type pollen (Figure 3F). The altered
composition of the mature mutant pollen grains compared with
wild-type pollen, together with the undulations of the plasma
membrane, suggest a defect in the regulation of the plasma
membrane surface in the mutant pollen. The failure of mutant
TPLATE pollen to germinate is most likely caused by the ectopic
callose deposition between the plasma membrane and the
intine layer.
TPLATE-GFP Accumulates at the Pollen Tube Exit Site
upon Germination
Because pollen development requires TPLATE, we determined
the localization of TPLATE-GFP in germinating pollen. The
TPLATE-GFP protein was expressed in pollen from the Lat52
promoter, which is reported to have a pollen-specific expression
pattern with only low transcript levels detectable in anther walls
and petals (Ursin et al., 1989; Twell et al., 1990). Transcripts
driven by the Lat52 promoter are first detected in spores undergoing pollen mitosis I and then increase substantially in mature
pollen. GFP-tagged TPLATE protein expressed from Lat52
complemented the pollen phenotype of the tplate mutant. The
ratio of wild-type versus mutant pollen was examined in tplate
plants transformed with the Lat52-TPLATE-GFP construct. The
plants were heterozygous for both T-DNAs, so complementation
of the mutation caused by the insertion of the T-DNA in the
TPLATE gene should result in a shift from a 1:1 ratio of wild-type
versus mutant pollen to a 3:1 ratio. From 1325 pollen grains
counted, 971 had a wild-type appearance and 354 were shriveled. This amounts statistically to the expected 3:1 ratio of two
independently segregating insertions (x2 test for a 3:1 ratio yields
2.08, and x2 95% interval value is 3.84). The complementation
was further substantiated by wild-type Columbia (Col-0) backcrosses. The TPLATE T-DNA insertion was transmitted to the
wild-type plants via pollen that invariably also carried the
TPLATE-GFP gene. Moreover, using PCR, we identified plants
homozygous for the TPLATE T-DNA insertion in the backcrossed
offspring (data not shown). We conclude that the GFP-tagged
TPLATE protein, expressed by the Lat52 promoter, is functional
in pollen.
In mature pollen, TPLATE-GFP, expressed from the Lat52 or
the endogenous promoter, was cytoplasmic and granular (Figure
4A). Upon incubation of the pollen in germination medium,
TPLATE-GFP accumulated at a distinct area at the cell periphery
(Figure 4B). This region corresponds to the position where the
pollen tube emerges (Figure 4C). Time-lapse recording further
demonstrated that TPLATE-GFP is transported into the growing
pollen tube (Figures 4C and 4D). The localization of TPLATE-GFP
at the pollen tube exit site and at the tip of the pollen tube
supports a role in the control of vesicle fusion.
Expression Analysis of TPLATE
The pollen-maturation phenotype of TPLATE T-DNA insertion
plants and the subcellular localization of TPLATE-GFP in pollen
suggest that the corresponding gene is activated in developing
pollen. We analyzed the expression of TPLATE using a genomic
fragment containing the open reading frame and 800 bp upstream of the start codon fused to the b-glucuronidase gene.
b-Glucuronidase activity was detected in pollen grains and at the
connectivum, where the anther is connected to the filament (see
Supplemental Figure 1A online). A thorough study of wholegenome gene expression during different developmental stages
of Arabidopsis pollen was performed previously (Honys and
Twell, 2004). The TPLATE mRNA is expressed in uninucleate
microspores and in bicellular and immature tricellular pollen, but
it is low-abundant in dried, mature pollen grains. The same study
Cytokinesis and Pollen Depend on TPLATE
3507
Figure 3. Electron Microscopy of tplate and Wild-Type Pollen.
(A) Overview of mutant and wild-type pollen grains.
(B) Close-up of (A). Accumulation of callose is comparable to that seen in Figures 2G to 2I.
(C) Noncollapsed mutant pollen showing undulations of the plasma membrane (arrows). Bars ¼ 2 mm for (A) to (C).
(D) and (F) Immunoelectron microscopy sections of a mutant and a wild-type pollen grain using a callose-specific antibody. The mutant pollen shows
gold labeling between the plasma membrane (PM) and the intine layer (Int), whereas hardly any label is present in nongerminated wild-type pollen. Bar ¼
1 mm.
(E), (G), and (H) High-magnification images of wild-type and mutant pollen. Bars ¼ 1 mm.
pv, peripheral band of vesicles; v, vacuole.
showed that TPLATE expression is not restricted to pollen
(Honys and Twell, 2004) (see Supplemental Figure 1B online).
TPLATE was originally identified through homology with a tobacco BY-2 cDNA-AFLP tag (BSTT43-4-330) with an M-phase–
specific upregulation of expression (Breyne et al., 2002; Van
Damme et al., 2004a). We determined TPLATE mRNA levels in
Arabidopsis roots by whole-mount in situ hybridization. Transcript was present throughout the root tip meristem, although it
was not distributed uniformly. The lowest level of expression was
at the very tip end, and a stronger expression occurred near the
elongation zone (see Supplemental Figures 1C to 1F online). The
TPLATE expression pattern is in agreement with the results of
previous digital in situ results (Birnbaum et al., 2003; www.
arexdb.com) and does not match the expression pattern of cell
cycle–controlled genes. Thus, Arabidopsis TPLATE is not likely
to be cell cycle–controlled.
3508
The Plant Cell
TPLATE Is Required for Somatic Cytokinesis
Figure 4. TPLATE-GFP Localization in Pollen.
(A) Confocal section of a nongerminating mature pollen grain expressing
the TPLATE genomic construct showing diffuse labeling of TPLATEGFP. The appearance of signal near the periphery is predominantly
autofluorescent.
(B) Confocal section of TPLATE genomic-GFP accumulation at the future
pollen tube exit site.
(C) Confocal sections of a Z-stack (10 sections, 7.35 mm) time-lapse
The presence of TPLATE transcripts in somatic tissue suggests a
role for TPLATE in somatic processes besides pollen development. Arabidopsis lines homozygous for the TPLATE T-DNA
insertion that also carried the complementing Lat52-TPLATEGFP did not show discernible growth or a morphological phenotype. Because of Lat52-TPLATE-GFP expression in seedlings,
as shown by GFP fluorescence and protein gel blot analysis (see
Supplemental Figure 2 online), any somatic phenotype is likely to
be complemented in the homozygous TPLATE mutant lines by
TPLATE-GFP expression.
Therefore, we decided to suppress TPLATE activity by posttranscriptional gene silencing. The TPLATE cDNA was expressed
from the 35S promoter as a hairpin loop double-stranded RNA in
Arabidopsis. Two independent hairpin constructs with different
backbones yielded transgenic plants that displayed severe
growth defects (Figures 5A and 5B). The expression of TPLATE
in these plants was assessed by RT-PCR. Compared with wildtype plants, TPLATE expression was strongly reduced in the
knockdown seedlings (Figure 5C). The mRNA level of a control
gene, eIF-4A-1, was not affected, indicating that the growth
defect in knockdown seedlings did not influence the expression
of this household gene. The knockdown seedlings produced
thickened cotyledons with an irregular surface and a reduced
number of stomata (Figures 5A and 5B; data not shown). The
hypocotyl was approximately twice as thick as wild-type hypocotyls, whereas the root diameter appeared normal (Figures 5I
and 5J). Most plants were arrested at an early stage of development, although some of them produced a first set of
leaves that did not fully expand and had a vitrified appearance
(Figure 5B).
At 5 d after germination, mitotic cells were no longer detected
by DAPI staining and growth was completely arrested (data not
shown). A morphological analysis of PI-stained seedlings revealed
the presence of interrupted cell walls and aberrant cell plates in
epidermal cotyledon cells (Figure 5H) as well as in other cell types,
but these were more difficult to analyze three-dimensionally.
Cellulose polymerization defects and weakening of the cell wall
frequently go hand in hand with the stimulation of callose synthesis, which, besides its role in pollen development, normally
occurs only in wounded tissue and in developing cell plates
(Delmer and Amor, 1995; Nickle and Meinke, 1998; Gillmor et al.,
2005). To test the presence of ectopic callose deposition in RNA
interference (RNAi) Arabidopsis plants, transgenic material was
stained with aniline blue and epifluorescence was imaged. Numerous depositions of ectopic callose were detected in cotyledons (Figure 5F) and in roots (Figure 5K). In wild-type plants,
callose is detected in newly formed cell plates and in the junctions
of stomatal guard cells (Figures 5G and 5L). In RNAi seedlings,
callose depositions were most prominent in root tissue, in particular at the root tip (Figure 5K). The ectopic deposition of callose
series of germinating pollen grains showing accumulation of TPLATEGFP at the future pollen tube exit site.
(D) Punctate localization of TPLATE genomic-GFP at the pollen tube tip.
Cytokinesis and Pollen Depend on TPLATE
3509
Figure 5. Silencing of the TPLATE Gene in Arabidopsis Seedlings.
(A) Overview of a TPLATE knockdown and a wild-type seedling germinated without selection.
(B) RNAi seedling forming a first pair of true leaves.
(C) Agarose gel showing reduced amplification of the TPLATE mRNA in the knockdown seedlings and amplification of eIF-4A-1 as a control (left lanes)
and amplification of both TPLATE and eIF-4A-1 in a single reaction (right lanes).
(D) and (E) Confocal Z-stack projections taken at the same magnification of the root tip of an RNAi and a wild-type seedling stained with FM4-64. In RNAi
mutants, root hairs emerge close to the tip (white arrowhead). The inset in (D) shows the aberrant morphology of the root tip in these RNAi seedlings.
(F) and (G) Epifluorescence images of an RNAi and a wild-type cotyledon showing ectopic callose deposition (yellow) visualized by aniline blue staining.
The inset in (F) is a blow-up of an epidermal cell showing depositions of callose. Red signal is autofluorescent. In wild-type cotyledons, aniline blue
labels only newly formed cell plates and stomatal guard cells (inset in [G]).
(H) Confocal section of PI-stained epidermal cells of TPLATE knockdown seedlings showing incomplete cell plates. Bars ¼ 10 mm.
(I) and (J) Differential interference contrast images of the hypocotyl–root interphase (arrowheads) of an RNAi and a wild-type seedling.
(K) and (L) Confocal sections of an RNAi and a wild-type root stained with aniline blue. Images were made using the exact settings of laser power and
pinhole diameter to visualize callose. Callose accumulates strongly in RNAi seedling roots compared with wild-type roots, where callose is present only
in forming cell plates. Bars ¼ 20 mm.
in silenced plants and cell cultures is indicative of a malfunctioning of cell wall formation or modification.
To investigate the growth inhibition and cytokinesis defects
observed in Arabidopsis in more detail, TPLATE was silenced in
BY-2 tobacco suspension cells using a homologous cDNA-AFLP
fragment previously isolated from these cells (tag BSTT43-4330) (Breyne et al., 2002). The TPLATE RNAi calli grew significantly slower than calli transformed with unrelated constructs
(data not shown). In >20 independent transformation events that
were analyzed, ;10% of the BY-2 cells had misplaced cell walls
3510
The Plant Cell
(deviations $ 908) or produced cell walls with severe deformations. Examples of aberrant cross walls are presented in Supplemental Figure 3 online.
In addition, we observed that cell plates did not always
fuse to the mother wall and produced fuzzy extreme ends. Figure
6 shows a time-lapse recording of a TPLATE RNAi BY-2 cell
stained with FM4-64 that failed to anchor its newly formed cell
plate.
The plate is initially made and expands until it reaches the
cortex (Figure 6, image after 62 min) but subsequently fails to
insert, even when the cell is followed for >90 min after contact of
the cell plate with the cortex. The cell in Figure 6 also shows
ectopic growth, which is commonly observed in TPLATE RNAi
BY-2 cell lines and may be caused by mistargeting of cell wall–
modifying factors, as a result of the reduced activity of TPLATE.
The RNAi effects in Arabidopsis and BY-2 cells confirm that
TPLATE is required for somatic cytokinesis and anchoring of the
cell plate with the mother wall.
Root and Hypocotyl Tissues from TPLATE RNAi Plants
Are Severely Disorganized
Root morphology was determined for several RNAi plants (Figures 5D and 5E). The meristematic tissue contained differentiated cells, and root hairs emerged close to the tip (Figure 5D,
arrowhead). Figure 7 shows toluidine blue–stained transverse
sections of wild-type and TPLATE RNAi root–hypocotyl and
hypocotyl sections.
Some radial patterning can be observed in the RNAi seedlings,
with smaller cells in the vascular bundle surrounded by larger
cells of the endodermis, cortex, and epidermis. However, the
vascular cylinder in the RNAi plants is severely disorganized, and
cortical layers cannot be distinguished clearly (Figure 7B). In all
transverse TPLATE RNAi hypocotyl sections examined, the cyan
blue coloration typical for toluidine blue–stained wild-type xylem
vessel cells (arrows in Figure 7A) was absent, indicating that
these seedlings did not produce differentiated xylem elements.
Figures 7C and 7D show sections at identical magnifications
through the hypocotyl of a wild-type and an RNAi seedling. The
central cylinder and cortical layers are present in the RNAi
seedling section, although it is difficult to discriminate between
the different layers. Incomplete and malformed cell walls are
observed in the cortex of RNAi hypocotyls (Figures 7D, inset, and
7E). In addition, there are extra random divisions, predominantly
in the vascular tissue (Figure 7D), leading to an increase in
hypocotyl diameter. The central cylinder cells are not converted
to vascular tissue, as in the wild type. Instead, they differentiate
to starch-containing cells, as revealed by Lugol staining (Figure
7F, inset). Starch accumulation inside the vascular bundle was
never observed in wild-type hypocotyl sections (data not shown).
Therefore, reducing TPLATE expression leads to altered differentiation and extra rounds of cell division, predominantly in the
vascular bundle.
Figure 6. RNAi Effects in BY-2 Time Lapse.
Time-lapse recording of a BY-2 cell transformed with the pH7GWIWG(II) vector containing the tobacco TPLATE tag BSTT43-4-330, showing a dividing
cell stained with FM4-64 that fails to insert the newly formed cell plate. Bars ¼ 20 mm.
Cytokinesis and Pollen Depend on TPLATE
3511
TPLATE Connects the Cell Plate with the Mother Wall
To investigate the subcellular localization of TPLATE in a tissue
context, we analyzed Arabidopsis plants carrying an N- or
C-terminally fused TPLATE construct with GFP behind the 35S
promoter. Arabidopsis leaves were imaged with the confocal
microscope within the first 24 to 36 h after germination, when
numerous leaf epidermal cells and root meristematic cells divide
(Figure 8).
GFP fluorescence was observed mainly in the cytoplasm
of nondividing cells with both constructs. During cytokinesis,
N- and C-terminally fused TPLATE concentrated at the developing cell plate (Figure 8, asterisks). The fusion proteins associated
with the newly formed cell plate at an early stage of phragmoplast
development (Figures 8C, 8D, and 8I). Upon contact of the cell
plate with the mother wall, GFP fluorescence accumulated at the
contact site, spreading over a region of ;5 mm surrounding the insertion site (Figures 8E and 8F, yellow arrowheads, and inset in 8F).
The accumulation of TPLATE fused to GFP at the division zone,
together with the observation that in RNAi lines cell plates failed
to or did not correctly connect to the mother wall, suggest that
TPLATE plays a role in anchoring the cell plate to the mother wall.
DISCUSSION
Figure 7. Transverse Hypocotyl Sections of RNAi Seedlings Reveal
Aberrant Cell Fates of Vascular Cells.
(A) Toluidine blue–stained section of a wild-type root–hypocotyl junction
and blow-up of the vascular system. Xylem cells are stained light blue
and indicated with arrows.
(B) Toluidine blue–stained section of an RNAi root–hypocotyl junction
and blow-up of the vascular system. Small cells in the vascular cylinder
are formed, but based on the absence of the blue coloration representative of xylem cells, differentiation and patterning are lost.
(C) Toluidine blue–stained wild-type hypocotyl section.
(D) Toluidine blue–stained RNAi hypocotyl section taken at the same
magnification. Arrows indicate incomplete cell plates. The inset shows a
blow-up of a cell with aberrant division plane orientation.
(E) Blow-up of the vascular system of an RNAi hypocotyl section. Arrows
indicate misguided cell plates.
(F) RNAi hypocotyl section from Figure 6D and blow-up (inset) stained
with Lugol. Vascular cells accumulate starch granules.
Bars ¼ 100 mm.
TPLATE is a plant-specific protein that is essential for the formation of viable pollen grains and for the final steps of cytokinesis in
somatic cells. This conclusion is supported by the male sterility of
a tplate T-DNA insertion mutant, by the observation that TPLATEGFP temporally accumulated at a narrow zone along the plasma
membrane where the cell plate makes contact with the mother
wall, and by the observation that downregulation of TPLATE
results in anchoring defects. Nonconventional types of cytokinesis, such as male and female gametophyte development, were
apparently unaffected in the tplate mutant, suggesting that the
cortical localization of TPLATE-GFP is driven by a PPB-dependent process. In agreement with this notion, the peripheral region
occupied by TPLATE-GFP corresponds to the division zone
formerly marked by the PPB during the early steps of mitosis. The
accumulation of TPLATE when the cell plate is in close proximity
to the mother wall suggests that TPLATE contributes to cell plate
formation and positioning during the final steps of cytokinesis.
Pollen germination and somatic cytokinesis involve substantial
vesicle transport and fusion and, therefore, not surprisingly,
become rate-limiting when one of the components is reduced or
missing. Arabidopsis mutants with defective vesicle transport
machinery usually display pleiotropic cellular phenotypes that
culminate in a general growth disorder (Kang et al., 2001;
Sanderfoot et al., 2001; Geelen et al., 2002). However, vesicle
trafficking and fusion can also be dependent on a highly selective
component that is restricted to a single process. The prime
example for such selectivity is the KNOLLE syntaxin specifically
involved in somatic cytokinesis (Müller et al., 2003).
Here, we describe a putative membrane traffic protein that is
required in two seemingly unrelated cellular processes: pollen
germination and somatic cytokinesis. Both phenomena necessitate the modification of the cell wall at a specific site where new
wall material joins the existing wall. The targeting of GFP-tagged
3512
The Plant Cell
Figure 8. Localization of TPLATE-GFP and GFP-TPLATE in Arabidopsis Root and Leaf Epidermal Cells.
(A) and (B) Overview of TPLATE-GFP and GFP-TPLATE in Arabidopsis root cells. The fusion protein accumulates in the midline of dividing cells
(asterisks).
(C) and (D) Close-up of midline targeting of the fusion protein in a young phragmoplast of Arabidopsis root cells (between white arrowheads).
(E) and (F) Close-up of the fusion protein accumulating in the midline of the phragmoplast and the division zone (yellow arrowheads). The inset in (E)
shows that plasma membrane labeling of TPLATE-GFP is absent before plate insertion (asterisk) into the mother wall. The inset in (F) shows a blow-up
of the accumulation of GFP-TPLATE (yellow bar) at the division zone upon plate insertion.
(G) and (H) Localization of GFP-TPLATE and TPLATE-GFP in leaf epidermal cells of Arabidopsis at 24 to 36 h after germination. Asterisks indicate
accumulation of the fusion protein in the midline of dividing cells.
(I) Magnification of a dividing epidermal cell (indicated with I in [H]) showing midline targeting of TPLATE-GFP.
TPLATE during cytokinesis and pollen germination to the division
site and the pollen tube exit site, respectively, suggests that
TPLATE contributes to these cell wall modifications (see Supplemental Figure 4 online).
cytokinetic cell (Jürgens, 2005a). Similar to TPLATE, COPI proteins accumulate at the phragmoplast midline, where the cell
plate is formed (Couchy et al., 2003; Vanstraelen et al., 2006). To
date, it is not yet clear how important these coat proteins are for
the formation of the cell plate.
The TPLATE Protein Shows Similarity to the COPI Proteins
Adaptor protein complexes are involved in the formation of
clathrin-coated vesicles, occurring in endocytic and recycling
pathways between the plasma membrane and the lysosomes or
the trans-Golgi network (Kirchhausen et al., 1997; McMahon and
Mills, 2004). The TPLATE protein shows similarity to the adaptin_N
domain of COP proteins; in addition, TPLATE proteins contain a
14–amino acid motif that is highly conserved in all b-COP
proteins. We designated this motif the b-COP–specific element.
Cell plate formation relies on massive vesicle transport along
the phragmoplast microtubules and involves homotypic fusion of
these vesicles to produce a solid cell plate at the center of the
TPLATE-GFP Concentrates at Sites Where Vesicle Fusion
Takes Place
GFP-tagged TPLATE concentrates between the separated chromatin when the phragmoplast emerges at the cell center. The
GFP signal follows the laterally expanding cell plate up to the
moment of contact with the mother wall (Van Damme et al.,
2004a). Remarkably, TPLATE also concentrates at the cortical
division zone when the cell plate approaches the mother wall
(Figures 8E and 8F; see Supplemental Figure 4 online). The
protein accumulates along a stretch ;5 mm wide at the final
insertion site.
Cytokinesis and Pollen Depend on TPLATE
At the cell plate, vesicle fusion drives cell plate formation and
allows the plate to expand in a centrifugal manner. The vesicle
fusion process is mediated by an essential, cytokinesis-specific
syntaxin (KNOLLE), a SNARE member that cooperates with other
membrane proteins to form a vesicle fusion complex (Lauber
et al., 1997; Müller et al., 2003; Jürgens, 2005a). Because cell
plates are made de novo, at first there is no target membrane and
vesicle fusion must primarily be homotypic. Plasma membrane
and cell plate composition are not identical; thus, attachment of
the cell plate to the mother wall probably depends on a heterotypic fusion process that may require a different set of SNARE
proteins (Jürgens, 2005a; Vanstraelen et al., 2006). knolle mutants are impeded in making correct cell plates but frequently
produce small cell wall stubs associated with the mother wall.
These stubs may result from continued vesicle deposition at the
insertion site in an attempt to make contact with a cell plate that
in the end does not form. The formation of these stubs in the
knolle mutant indicates that the KNOLLE complex is not essential
for the connection of the cell plate with the mother wall. A cell
plate was still formed in TPLATE knockdown Arabidopsis cells
and BY-2 cell lines. However, these plates were not rigid as in
wild-type cells, and they did not always connect properly to the
mother wall. Possibly, cell plates did not attach correctly to the
mother wall and therefore did not mature as in wild-type cells,
leading to crooked shapes, in agreement with previous reports
(Palevitz, 1980; Mineyuki and Gunning, 1990).
In germinating pollen, TPLATE-GFP accumulates in a punctate
manner at the site of pollen tube exit and later travels toward the
tip of the growing pollen tube (Figure 4; see Supplemental Figure
4 online). Pollen tube growth relies heavily on extensive vesicle
transport to the tube tip and subsequent fusion of vesicles to the
plasma membrane to allow expansion and the secretion of cell
wall components (Hepler et al., 2001). The secretion of vesicles is
a heterotypic fusion process whereby specific target SNAREs or
syntaxins at the plasma membrane are needed (Carter et al.,
2004). The involvement of TPLATE in pollen tube emergence and
growth distinguishes it from KNOLLE, because the knolle mutants do not produce defective pollen grains. The function of
KNOLLE is restricted to homotypic vesicle fusion processes that
may not be primordial for pollen tube formation (Müller et al.,
2003). TPLATE, on the other hand, might be important in facilitating heterotypic vesicle fusion.
Ectopic Callose Deposition in tplate Mutants
Mature tplate mutant pollen contained extensive depositions of
callose laid as irregular formations against the intine cell wall
layer. Intine and exine appeared morphologically identical to
those of wild-type pollen, indicating that callose deposition did
not interfere with cell wall formation and likely occurred after wall
formation had been completed. Based on appearance in electron micrographic recordings, differences in vesicle composition
between wild-type and mutant pollen were observed. The most
affected pollen contained fewer vesicles, and the diversity based
on electron density was altered. Whether these differences in
vesicle composition are the cause of ectopic callose formation
remains to be determined. Relatively little is known about the
composition and dynamics of vesicles in pollen. Most informa-
3513
tion comes from electron microscopic observations, by which it
was determined that cell wall precursor–containing vesicles
localize predominantly at the peripheral border near the intine
layer (Van Aelst et al., 1993). The precursor vesicles accumulate
in large amounts at the periphery to support the very fast mode of
germination observed for Arabidopsis pollen (Heslop-Harrison,
1987; Pickert, 1988). Because TPLATE-GFP is concentrated at
the cell periphery during pollen exit site establishment and at the
pollen tube tip, it is possible that TPLATE is involved in vesicle
targeting and fusion with the plasma membrane.
The TPLATE pollen morphology strongly resembles that of
adl1C mutant pollen (Kang et al., 2003a; ADL1C was renamed
DRP1C by Hong et al. [2003a]). The adl1C mutation specifically
affects male gametogenesis and leads to the production of
shriveled pollen that fails to germinate. Remarkably, the adl1C
and tplate mutations are fully penetrant, which is not so for
several other male-sterile mutants (Kang et al., 2003a; this work).
Similar to tplate pollen, adl1C pollen grains contain cell wall
depositions close to the intine layer, the plasma membrane is
heavily undulated, and vesicle composition is changed. The
desiccation intolerance and shriveled appearance of both adl1C
and tplate pollen is in agreement with a critical requirement of an
intact intine wall for pollen germination and survival (Grini et al.,
1999; Fei and Sawhney, 2001). DRP1C is a member of the
dynamin-like protein family, which in animal cells function in both
clathrin-mediated and non-clathrin-mediated endocytic processes (Praefcke and McMahon, 2004).
Plant dynamins are subdivided into six families based on
functional motifs (Hong et al., 2003a). Members of the Arabidopsis DRP1 family, which is most related to soybean (Glycine max)
PHRAGMOPLASTIN, mediate membrane tubule formation and
function in vesicle trafficking from Golgi to the cell plate (Gu and
Verma, 1996; Hong et al., 2003b; Kang et al., 2003a, 2003b). The
striking similarity between the pollen phenotypes conferred by
tplate and adl1C together with the cell plate localization of both
proteins during cytokinesis reinforce the idea that TPLATE functions in the regulation of vesicular traffic. Callose depositions
were also observed in dividing cells in cotyledons and in roots in
TPLATE RNAi seedlings, where vesicle composition is likely very
different from that in pollen.
Ectopic callose is often regarded as an indication of compromised cell wall integrity and occurs in response to stress situations (Gillmor et al., 2005). Defective cellulose deposition
triggered by cellulose biosynthesis inhibitors, such as dichlorobenzonitrile or isoxaben (Desprez et al., 2002), or attributable to
a cellulose deficiency mutation, as in cyt1 embryos (Nickle and
Meinke, 1998; Lukowitz et al., 2001), leads to the accumulation of
ectopic callose. Ectopic callose has also been seen to accumulate in embryos carrying a double mutation in the dynamins adl1A
and adl1E (Kang et al., 2003b). The overexpression of PHRAGMOPLASTIN or a GTPase-defective form of PHRAGMOPLASTIN
caused a persistence of callose in fully completed cell plates.
Here, the mispositioning of cell plates during megaspore development is believed to retard or obstruct cell plate maturation
(Geisler-Lee et al., 2002; Hong et al., 2003b). Cell plates that
connect with the mother wall at unprepared sites seem not to
flatten (Palevitz, 1980), possibly because the division zone contains some essential cell wall maturation factors (Mineyuki and
3514
The Plant Cell
Gunning, 1990). The embryonal cytokinesis defects in knolle and
keule mutants do not display the accumulation of ectopic callose
(Nickle and Meinke, 1998). Incomplete cell wall formation in
plants without callose accumulation is phenocopied by caffeine,
an inhibitor of Ca2þ-controlled vesicle-trafficking processes
(Hepler and Bonsignore, 1990) that resembles the cyd mutant
of pea (Pisum sativum) (Liu et al., 1995). It is not the intrinsic
strength of the cell wall in these cytokinesis mutants that is
affected; rather, walls are missing or incomplete. The observations that cell plates can form and that callose persists in TPLATE
RNAi seedlings and BY-2 cells suggest that TPLATE is more
likely to be involved in cell wall modification and maturation than
in the primary steps of plate formation.
TPLATE Is Required for Cell Plate Positioning
and Cell Differentiation
Reduced expression of the TPLATE gene by RNAi resulted in the
formation of distorted and sometimes incomplete cell walls.
TPLATE is a singleton in Arabidopsis with low overall nucleotide
similarity to genomic sequences, reducing the chance that other
genes are silenced too. Knockdown plants sometimes produced
a first pair of leaves. Regardless of the development of leaves, all
plants died within a few weeks after germination. It is possible
that the extended life span compared with other cytokinesis
mutants is attributable to the low activity of the 35S promoter
during embryogenesis, allowing growth beyond the embryonic
heart stage (Odell et al., 1994; Völker et al., 2001).
TPLATE RNAi seedlings had a radially expanded hypocotyl
similar to that of the cellulose-deficient mutants kor1-2/rsw2,
rsw1/cesA1, kob1, and procuste1 (Arioli et al., 1998; Nicol et al.,
1998; Fagard et al., 2000; Peng et al., 2000; Lane et al., 2001;
Williamson et al., 2001; Beeckman et al., 2002; Pagant et al.,
2002). With the exception of kor1-2, these mutants have a thickened hypocotyl without severe impact on tissue organization or
development of the vascular tissue. The kor1-2 mutant lacks
distinct cell files in roots, hypocotyls, and cotyledons, shows
impaired vascular tissue development, and cell divisions occur
randomly (Zuo et al., 2000). Similar defects in cell file organization
and random cell divisions are found in the TPLATE RNAi seedlings. At the cellular level, kor1-2 mutants show misplaced and
distorted cell plates that often terminate before reaching the
mother wall (Zuo et al., 2000). TPLATE RNAi seedlings also
produced cells with interrupted and misplaced cell walls. The
imperfect separation of dividing cells may account for miscommunication among neighboring cells whereby cell differentiation
is disturbed and the tissues become improperly organized
(Lukowitz et al., 1996). Perhaps such miscommunication provokes the formation of the starch-containing cells seen in the
TPLATE RNAi hypocotyl sections. The occurrence of inadvertent
cell divisions at the central cylinder is probably not responsible
for the altered differentiation in the hypocotyl of TPLATE RNAi
seedlings, as additional vascular divisions also occur in the
mutant fass without a loss of correct vascular differentiation
(Torrez-Ruiz and Jürgens, 1994). Extra cell divisions in the central
cylinder have also been reported for the cellulose-deficient
rsw2-1 mutant of Arabidopsis, which is allelic to kor1-2 (Lane
et al., 2001). KOR1 encodes a membrane-anchored endogluca-
nase (KORRIGAN) that is assumed to function in cell wall modification and maturation. TPLATE may contribute to cell wall
maturation too, as cell plates and cell walls appear less strong in
RNAi BY-2 cells.
Although several lines of evidence support a direct role for
TPLATE in cell plate maturation, it is possible that incomplete cell
walls and ectopic callose deposition are secondary effects of
TPLATE silencing. Nevertheless, the subcellular distribution of
TPLATE holds the promise that its primary function is to ascertain
correct plasma membrane targeting (see Supplemental Figure 4
online). In the absence of TPLATE, specialized vesicles are no
longer correctly formed or, more likely, they are inapt for sitedirected targeting to the plasma membrane. The fact that the
TPLATE mutation was pronounced in pollen and did not have a
discernible effect on female gametophyte fertility may reflect a
differential requirement for plasma membrane targeting between
these cell types. TPLATE is clearly also essential for normal
somatic development. Silencing of TPLATE has pleiotropic effects, including changes in cell division and cellular differentiation. Together, our findings have led to an unexpected link
between the vesicle–plasma membrane interphase in pollen and
cell plate maturation and attachment during cytokinesis.
METHODS
T-DNA Insertion Mutant Analysis
Arabidopsis thaliana SALK_0030086 was obtained from the SALK collection (Alonso et al., 2003). Genomic DNA was extracted, and PCR
to identify the insertion was done using the LP and RP primers
(59-CCCTTGGCAATTTTCCTCGAA-39 and 59-TGCAAAGTGACCCTTCCATCA-39) that were recommended by the SIGNaL iSect Tool of The
Arabidopsis Information Resource database (www.Arabidopsis.org) in
combination with the T-DNA–specific LBa1 primer (59-TGGTTCACGTAGTGGGCCATCG-39).
Thin Sections
Whole-mount anther and seedling sections were prepared according to
De Smet et al. (2004). Thin sections (5 mm) were made using a Rotation
Microtome 2040 (Leica Microsystems) using glass knives (Leica Microsystems) for transverse sections and Superlap knives (Adamas Instrumenten) for longitudinal anther sections. Transverse sections of roots and
hypocotyls were stained with 0.05% toluidine blue O (Sigma-Aldrich) in
0.1 M NaPO4 buffer, pH 7.0, and 1% Lugol solution (Merck) for starch
staining.
Pollen Analysis
Pollen morphology was analyzed using light microscopy (DMLB; Leica).
The pollen was stained with Alexander stain (Alexander, 1969), and
mutant versus wild-type pollen grains were counted. Germination of
pollen grains in vitro was performed as described by Chen and
McCormick (1996). Anthers were stained with DAPI staining solution
(0.1% Nonidet P-40, 10% DMSO, 50 mM PIPES, 5 mM EGTA, and 1 mL of
DAPI; 1 mg/mL).
PI (Sigma-Aldrich) staining of thin anther sections was done by adding
30 mM PI to the slides and washing them after 5 min of incubation. Aniline
blue (0.5% in water; Sigma-Aldrich) and calcofluor white (1% in water,
fluorescent brightener 28; Sigma-Aldrich) were added simultaneously to
the thin anther sections, and slides were washed after 5 min of incubation.
Cytokinesis and Pollen Depend on TPLATE
Fluorescent Protein Fusion Constructs and Transformation
The construct to drive TPLATE-GFP expression under the control of the
Lat52 promoter was created using the multisite Gateway system (Invitrogen) (Karimi et al., 2005). pDONRP4-P1R-Lat52, pDONR207-TPLATE,
and pDONRP2R-P3-EGFP were used to clone the Lat52 promoter, the
TPLATE, and the EGFP open reading frames in pH7m34GW. pLAT52-7
(Bate et al., 1996) was kindly provided by David Twell and was used for
PCR of the Lat52 promoter and cloning in pDONRP4-P1R using the
following primer pair: 59-GGGGACAACTTTGTATAGAAAAGTTGGTCGACATACTCGACTCAGAAGGTAT-39 and 59-GGGGACTGCTTTTTTGTACAAACTTGGTGTCTTGTTTTGATTATA-39.
The genomic fragment of the TPLATE gene (5.1 kb) was amplified
by PCR using Gateway adapted primers (FW, 59-GGGGACAAGTTTGTACAAAAAAGCAGGCTTTTATTGTTCCTCACAGTCATGATATCGC-39;
REV, 59-GGGGACCACTTTGTACAAGAAAGCTGGGTTGTTAACTTTGGTATATTTTCTATCTTTGCAC-39) and cloned into an adapted version of
the pK7GWFS7 plasmid to allow in-frame translational fusion between
the TPLATE protein, b-glucuronidase, and EGFP. Expression of the
TPLATE messenger was analyzed by b-glucuronidase expression, and
the fusion protein was localized by GFP fluorescence using confocal
microscopy.
35S-GFP fusion constructs and transformation of tobacco (Nicotiana
tabacum) BY-2 cells have been described by Van Damme et al. (2004a,
2004b). Cotransformations in BY-2 cells were obtained by consecutive
transformations. Arabidopsis ecotype Col-0 plants were transformed by
Agrobacterium tumefaciens–mediated transfection using the floral dip
method (Clough and Bent, 1998).
Microscopy and Drug Treatments
Image acquisition was as described by Van Damme et al. (2004b). PI
staining and differential interference contrast images of anther sections
were obtained with a 100M confocal microscope with software package
LSM510 version 3.2 (Zeiss) equipped with a 633 water-corrected objective (numerical aperture ¼ 1.2) using the settings for RFP excitation and
emission (excitation ¼ 543 nm; emission ¼ 560 nm cutoff). Aniline blue,
calcofluor, and DAPI staining were imaged using an Axiovert 135M
fluorescence microscope equipped with a 633 water-corrected objective (numerical aperture ¼ 1.2) with emission and excitation filters
(excitation ¼ 395 nm; emission ¼ 420 nm long pass), and images were
taken with an Axiocam camera and Axiovision software version 3.1
(Zeiss). Thin root sections on slides were analyzed by a light microscope
(DMLB; Leica) equipped with a 403 objective, and images were taken
with an Axiocam camera and Axiovision software version 3.1 (Zeiss).
Arabidopsis seedlings were imaged between slide and cover glass.
Aniline blue (0.5% in water) and FM4-64 (50 mM; Invitrogen) were added
to whole Arabidopsis seedlings and imaged after a 5-min incubation.
BY-2 cells were applied to a chambered cover glass system (Lab-Tek).
Cells were immobilized in a thin layer of 200 mL of BY-2 medium containing vitamins and 0.8% low-melting-point agar (Invitrogen). FM4-64
(50 mM) and PI (30 mM) were added to 1 mL of liquid BY-2 medium with
vitamins, and the drug concentration was adjusted to a final volume of
1.2 mL before addition to the samples. Stock solutions of FM4-64 (5 mM)
and PI (1.5 mM) were prepared in nanopure water. Fluorescence intensity
graphs were made using the profile program of the LSM software (Zeiss).
Pollen expressing Lat52-TPLATE-GFP was analyzed by confocal microscopy on slides coated with germination medium.
3515
mixture; Duchefa), 10 g/L sucrose (Acros Organics), 100 mg/L myoinositol (Sigma-Aldrich), 0.5 g/L MES (Duchefa), pH 5.7, with KOH, and
8 g/L plant tissue culture agar (International Diagnostics Group) with
15 mg/mL hygromycin B (Duchefa). The presence of the T-DNA was
identified by PCR using the RP and LBaI primers. Pollen of T-DNA– and
Lat52-TPLATE-GFP–positive plants was used to pollinate Col-0 plants,
and transmission of the T-DNA insertion via the parental route was
checked by PCR on the offspring plants.
Electron Microscopy Analysis
For scanning electron microscopy, air-dried pollen was tapped on stubs,
sputter-coated with gold, and examined with a scanning microscope
(JEOL) under an acceleration voltage of 10 kV. For transmission electron
microscopy, tissue samples were immersed in a fixative solution of 2%
paraformaldehyde and 2.5% glutaraldehyde and postfixed in 1% OsO4
with 1.5% K3Fe(CN)6 in 0.1 M Na-cacodylate buffer, pH 7.2. Samples
were dehydrated through a graded ethanol series, including a bulk
staining with 2% uranyl acetate at the 50% ethanol step, followed by
embedding in Spurr’s resin. Ultrathin sections, made on an Ultracut E
microtome (Leica Microsystems), were poststained in an ultrastainer
(Leica) with uranyl acetate and lead citrate. Samples were viewed with a
1010 transmission electron microscope (JEOL).
For immunoelectron microscopy analysis, root tips of 4-d-old Arabidopsis seedlings (Col-0 ecotype) were excised, immersed in dextran
(20%), and frozen immediately in a high-pressure freezer (EM Pact; Leica
Microsystems). Freeze substitution was performed in a Leica EM AFS.
Over a period of 4 d, root tips were substituted in dry acetone þ 0.1%
OsO4. Samples were infiltrated at 48C stepwise in Spurr’s resin and
embedded in molds. The polymerization was performed at 708C for 16 h.
Ultrathin sections of gold interference color were cut using an ultramicrotome (ultracut E; Reichert-Jung) and collected on formvar copper slot
grids. All steps of immunolabeling were performed in a humid chamber at
room temperature. Samples were blocked in blocking solution (5% BSA
and 1% fish skin gelatin in PBS) for 15 min followed by a wash step for
5 min (1% BSA in PBS). Incubation in a dilution (1% BSA in PBS) of
primary antibodies (anti-callose 1:100; Biosupplies Australia) for 60 min
was followed by washing four times for 5 min each (0.1% BSA in PBS).
The grids were incubated with 10-nm Protein A Gold (Cell Biology,
Utrecht University) and washed twice for 5 min each (0.1% BSA in PBS,
PBS, and double distilled water). Sections were poststained in a LKB
ultrastainer for 30 min in uranyl acetate at 408C and for 5 min in lead stain
at 208C. Control experiments consisted of treating sections with 10-nm
Protein A Gold alone. Grids were viewed with a JEOL 1010 transmission
electron microscope operating system at 80 kV.
RNAi Experiments
The TPLATE coding sequence was cloned into the pK7GWIWG(II) or
pH7GWIWG(II) vector (Karimi et al., 2002) to produce double-stranded
RNA, and Arabidopsis Col-0 plants were transformed. Transformed seeds
were selected on K1 medium with 25 mg/mL kanamycin A (Duchefa) or
15 mg/mL hygromycin B (Duchefa). BY-2 cells were transformed with the
vector pH7GWIWG(II) containing the BSTT43-4-330 tobacco tag (Breyne
et al., 2002), and calli were selected on hygromycin B (30 mg/mL;
Duchefa).
RT-PCR
Complementation of the TPLATE Mutation
Plants heterozygous for the T-DNA insertion in TPLATE (SALK_0030086)
were transformed with the Lat52-TPLATE-EGFP construct. Seeds were
selected on K1 medium (4.308 g/L Murashige and Skoog basal salt
Arabidopsis seedlings were ground, and the RNA was extracted using the
RNeasy kit (Qiagen). The RNA concentration was adjusted, and 1 mg of
total RNA was used for cDNA amplification (Superscript RNA polymerase
III; Invitrogen). PCR was performed using the following primer pairs:
3516
The Plant Cell
for TPLATE amplification, 59-CGTGTGGATATACGGGTTGGATT-39 and
59-CGCCAAAGATCATCATACCC-39 to amplify a 600-bp band; and for
eIF-4A-1 (At3g13920), 59-ACGGAGACATGGACCAGAAC-39 and 59-GCTGAGTTGGGAGATCGAAG-39 to amplify a 150-bp band.
Whole-Mount in Situ Hybridization
Probe synthesis and whole-mount in situ hybridization were performed as
described by Friml et al. (2003) on Arabidopsis Col-0 seedlings and
Arabidopsis 35S-TPLATE-GFP–expressing seedlings. The sense and
antisense probe mix consisted of a mixture of four separate probes
(250, 400, 490, and 600 bp) directed against different positions of the
TPLATE messenger.
Accession Number
The Arabidopsis Genome Initiative locus identifier for TPLATE is
At3g01780.
Supplemental Data
The following materials are available in the online version of this article.
Supplemental Figure 1. Expression Data of TPLATE.
Supplemental Figure 2. Ectopic Expression of Lat52-TPLATE-GFP.
Supplemental Figure 3. RNAi Effects in BY-2 Cells.
Supplemental Figure 4. Model of TPLATE Function in Pollen Germination and in Somatic Cytokinesis.
ACKNOWLEDGMENTS
We thank Rita Van Den Driesche for scanning electron microscopy,
Hugh Dickinson and Heather Owen for help with the pollen morphology
phenotype, David Twell for the Lat52 promoter, Toshiyuki Nagata and
Yasunori Machida for suggesting the callose staining, and Martine De
Cock for layout. S.C. is indebted to the Institute for the Promotion of
Innovation by Science and Technology in Flanders for a predoctoral
fellowship. D.V.D. is a Postdoctoral Fellow and D.G. was a Postdoctoral
Researcher of the Research Foundation-Flanders.
Received January 6, 2006; revised October 5, 2006; accepted November
10, 2006; published December 22, 2006.
REFERENCES
Alexander, M.P. (1969). Differential staining of aborted and non-aborted
pollen. Stain Technol. 44, 117–122.
Alonso, J.M., et al. (2003). Genome-wide insertional mutagenesis of
Arabidopsis thaliana. Science 301, 653–657.
Arioli, T., et al. (1998). Molecular analysis of cellulose biosynthesis in
Arabidopsis. Science 279, 717–720.
Bate, N., Spurr, C., Foster, G.D., and Twell, D. (1996). Maturationspecific translational enhancement mediated by the 59-UTR of a late
pollen transcript. Plant J. 10, 613–623.
Beeckman, T., Przemeck, G.K.H., Stamatiou, G., Lau, R., Terryn, N.,
De Rycke, R., Inzé, D., and Berleth, T. (2002). Genetic complexity of
CESA gene function in Arabidopsis embryogenesis. Plant Physiol.
130, 1883–1893.
Birnbaum, K., Shasha, D.E., Wang, J.Y., Jung, J.W., Lambert, G.M.,
Galbraith, D.W., and Benfey, P.N. (2003). A gene expression map of
the Arabidopsis root. Science 302, 1956–1960.
Boehm, M., and Bonifacino, J.S. (2001). Adaptins: The final recount.
Mol. Biol. Cell 12, 2907–2920.
Breyne, P., Dreesen, R., Vandepoele, K., De Veylder, L., Van
Breusegem, F., Callewaert, L., Rombauts, S., Raes, J., Cannoot,
B., Engler, G., Inzé, D., and Zabeau, M. (2002). Transcriptome
analysis during cell division in plants. Proc. Natl. Acad. Sci. USA 99,
14825–14830.
Carter, C.J., Bednarek, S.Y., and Raikhel, N.V. (2004). Membrane
trafficking in plants: New discoveries and approaches. Curr. Opin.
Plant Biol. 7, 701–707.
Chen, Y.C., and McCormick, S. (1996). Sidecar pollen, an Arabidopsis
thaliana male gametophytic mutant with aberrant cell divisions during
pollen development. Development 122, 3243–3253.
Cleary, A.L., Gunning, B.E.S., Wasteneys, G.O., and Hepler, P.K.
(1992). Microtubule and F-actin dynamics at the division site in living
Tradescantia stamen hair cells. J. Cell Sci. 103, 977–988.
Clough, S.J., and Bent, A.F. (1998). Floral dip: A simplified method for
Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant
J. 16, 735–743.
Couchy, I., Bolte, S., Crosnier, M.-T., Brown, S., and SatiatJeunemaitre, B. (2003). Identification and localization of a b-COPlike protein involved in the morphodynamics of the plant Golgi
apparatus. J. Exp. Bot. 54, 2053–2063.
Delmer, D.P., and Amor, Y. (1995). Cellulose biosynthesis. Plant Cell 7,
987–1000.
De Smet, I., Chaerle, P., Vanneste, S., De Rycke, R., Inzé, D., and
Beeckman, T. (2004). An easy and versatile embedding method for
transverse sections. J. Microsc. (Oxf.) 213, 76–80.
Desprez, T., Vernhettes, S., Fagard, M., Refrégier, G., Desnos, T.,
Aletti, E., Py, N., Pelletier, S., and Hofte, H. (2002). Resistance
against herbicide isoxaben and cellulose deficiency caused by distinct mutations in same cellulose synthase isoform CESA6. Plant
Physiol. 128, 482–490.
Fagard, M., Desnos, T., Desprez, T., Goubet, F., Refregier, G.,
Mouille, G., McCann, M., Rayon, C., Vernhettes, S., and Höfte,
H. (2000). PROCUSTE1 encodes a cellulose synthase required for
normal cell elongation specifically in roots and dark-grown hypocotyls
of Arabidopsis. Plant Cell 12, 2409–2424.
Fei, H., and Sawhney, V.K. (2001). Ultrastructural characterization of
male sterile33 (ms33) mutant in Arabidopsis affected in pollen desiccation and maturation. Can. J. Bot. 79, 118–129.
Friml, J., Benkova, E., Mayer, U., Palme, K., and Muster, G. (2003).
Automated whole mount localisation techniques for plant seedlings.
Plant J. 34, 115–124.
Geelen, D., Leyman, B., Batoko, H., Di Sansebastiano, G.P., Moore,
I., and Blatt, M.R. (2002). The abscisic acid-related SNARE homolog
NtSyr1 contributes to secretion and growth: Evidence from competition with its cytosolic domain. Plant Cell 14, 387–406.
Geisler-Lee, C.J., Hong, Z., and Verma, D.P.S. (2002). Overexpression
of the cell plate-associated dynamin-like GTPase, phragmoplastin,
results in the accumulation of callose at the cell plate and arrest of
plant growth. Plant Sci. 163, 33–42.
Gillmor, C.S., Lukowitz, W., Brininstool, G., Sedbrook, J.C.,
Hamann, T., Poindexter, P., and Somerville, C. (2005). Glycosylphosphatidylinositol-anchored proteins are required for cell wall synthesis and morphogenesis in Arabidopsis. Plant Cell 17, 1128–1140.
Grini, P.E., Schnittger, A., Schwarz, H., Zimmermann, I., Schwab, B.,
Jürgens, G., and Hülskamp, M. (1999). Isolation of ethyl methanesulfonate-induced gametophytic mutants in Arabidopsis thaliana by a
segregation distortion assay using the multimarker chromosome 1.
Genetics 151, 849–863.
Cytokinesis and Pollen Depend on TPLATE
Gu, X., and Verma, D.P.S. (1996). Phragmoplastin, a dynamin-like
protein associated with cell plate formation in plants. EMBO J. 15,
695–704.
Hall, Q., and Cannon, M.C. (2002). The cell wall hydroxyproline-rich
glycoprotein RSH is essential for normal embryo development in
Arabidopsis. Plant Cell 14, 1161–1172.
Hepler, P.K., and Bonsignore, C.L. (1990). Caffeine inhibition of
cytokinesis: Ultrastructure of cell plate formation/degradation. Protoplasma 157, 182–192.
Hepler, P.K., Vidali, L., and Cheung, A.Y. (2001). Polarized cell growth
in higher plants. Annu. Rev. Cell Dev. Biol. 17, 159–187.
Heslop-Harrison, J. (1987). Pollen germination and pollen-tube growth.
Int. Rev. Cytol. 107, 1–78.
Hong, Z., Bednarek, S.Y., Blumwald, E., Hwang, I., Jurgens, G.,
Menzel, D., Osteryoung, K.W., Raikhel, N.V., Shinozaki, K.,
Tsutsumi, N., and Verma, D.P.S. (2003a). A unified nomenclature
for Arabidopsis dynamin-related large GTPases based on homology
and possible functions. Plant Mol. Biol. 53, 261–265.
Hong, Z., Geisler-Lee, C.J., Zhang, Z., and Verma, D.P.S. (2003b).
Phragmoplastin dynamics: Multiple forms, microtubule association
and their roles in cell plate formation in plants. Plant Mol. Biol. 53,
297–312.
Honys, D., and Twell, D. (2004). Transcriptome analysis of haploid male
gametophyte development in Arabidopsis. Genome Biol. 5, R85.1–
R85.13.
Jefferies, C.J., and Belcher, A.R. (1974). A fluorescent brightener used
for pollen tube identification in vivo. Stain Technol. 49, 199–202.
Jürgens, G. (2005a). Plant cytokinesis: Fission by fusion. Trends Cell
Biol. 15, 277–283.
Jürgens, G. (2005b). Cytokinesis in higher plants. Annu. Rev. Plant Biol.
56, 281–299.
Kang, B.-H., Busse, J.S., and Bednarek, S.Y. (2003a). Members of the
Arabidopsis dynamin-like gene family, ADL1, are essential for plant
cytokinesis and polarized cell growth. Plant Cell 15, 899–913.
Kang, B.-H., Busse, J.S., Dickey, C., Rancour, D.M., and Bednarek,
S.Y. (2001). The Arabidopsis cell plate-associated dynamin-like protein, ADL1Ap, is required for multiple stages of plant growth and
development. Plant Physiol. 126, 47–68.
Kang, B.-H., Rancour, D.M., and Bednarek, S.Y. (2003b). The dynaminlike protein ADL1C is essential for plasma membrane maintenance
during pollen maturation. Plant J. 35, 1–15.
Karimi, M., De Meyer, B., and Hilson, P. (2005). Modular cloning in
plant cells. Trends Plant Sci. 10, 103–105.
Karimi, M., Inzé, D., and Depicker, A. (2002). GATEWAY vectors for
Agrobacterium-mediated plant transformation. Trends Plant Sci. 7,
193–195.
Kirchhausen, T., Bonifacino, J.S., and Riezman, H. (1997). Linking
cargo to vesicle formation: Receptor tail interactions with coat proteins. Curr. Opin. Cell Biol. 9, 488–495.
Lane, D.R., et al. (2001). Temperature-sensitive alleles of RSW2 link the
KORRIGAN endo-1,4-b-glucanase to cellulose synthesis and cytokinesis in Arabidopsis. Plant Physiol. 126, 278–288.
Lauber, M.H., Waizenegger, I., Steinmann, T., Schwarz, H., Mayer,
U., Hwang, I., Lukowitz, W., and Jürgens, G. (1997). The Arabidopsis KNOLLE protein is a cytokinesis-specific syntaxin. J. Cell Biol. 139,
1485–1493.
Liu, C.-M., Johnson, S., and Wang, T.L. (1995). cyd, a mutant of pea
that alters embryo morphology is defective in cytokinesis. Dev. Genet.
16, 321–331.
Lloyd, C. (1995). Life on a different plane. Curr. Biol. 5, 1085–1087.
Lukowitz, W., Mayer, U., and Jürgens, G. (1996). Cytokinesis in the
Arabidopsis embryo involves the syntaxin-related KNOLLE gene
product. Cell 84, 61–71.
3517
Lukowitz, W., Nickle, T.C., Meinke, D.W., Last, R.L., Conklin, P.L.,
and Somerville, C.R. (2001). Arabidopsis cyt1 mutants are deficient
in a mannose-1-phosphate guanylyltransferase and point to a requirement of N-linked glycosylation for cellulose biosynthesis. Proc.
Natl. Acad. Sci. USA 98, 2262–2267.
McMahon, H.T., and Mills, I.G. (2004). COP and clathrin-coated vesicle
budding: Different pathways, common approaches. Curr. Opin. Cell
Biol. 16, 379–391.
Mineyuki, B., and Gunning, B.E.S. (1990). A role for preprophase
bands of microtubules in maturation of new cell walls, and a general
proposal on the function of preprophase band sites in cell division in
higher plants. J. Cell Sci. 97, 527–537.
Mineyuki, Y. (1999). The preprophase band of microtubules: Its function
as a cytokinetic apparatus in higher plants. Int. Rev. Cytol. 187, 1–50.
Müller, I., Wagner, W., Völker, A., Schellmann, S., Nacry, P., Küttner,
F., Schwarz-Sommer, Z., Mayer, U., and Jürgens, G. (2003).
Syntaxin specificity of cytokinesis in Arabidopsis. Nat. Cell Biol. 5,
531–534.
Nickle, T.C., and Meinke, D.W. (1998). A cytokinesis-defective mutant
of Arabidopsis (cyt1) characterized by embryonic lethality, incomplete
cell walls, and excessive callose accumulation. Plant J. 15, 321–332.
Nicol, F., His, I., Jauneau, A., Vernhettes, S., Canut, H., and Höfte, H.
(1998). A plasma membrane-bound putative endo-1,4-b-D-glucanase
is required for normal wall assembly and cell elongation in Arabidopsis. EMBO J. 17, 5563–5576.
Odell, J.T., Hoopes, J.L., and Vermerris, W. (1994). Seed-specific
gene activation mediated by the Cre/lox site-specific recombination
system. Plant Physiol. 106, 447–458.
Pagant, S., Bichet, A., Sugimoto, K., Lerouxel, O., Desprez, T.,
McCann, M., Lerouge, P., Vernhettes, S., and Höfte, H. (2002).
KOBITO1 encodes a novel plasma membrane protein necessary for
normal synthesis of cellulose during cell expansion in Arabidopsis.
Plant Cell 14, 2001–2013.
Palevitz, B.A. (1980). Comparative effects of phalloidin and cytochalasin-B on motility and morphogenesis in Allium. Can. J. Bot. 58,
773–785.
Peng, L., Hocart, C.H., Redmond, J.W., and Williamson, R.E. (2000).
Fractionation of carbohydrates in Arabidopsis root cell walls shows
that three radial swelling loci are specifically involved in cellulose
production. Planta 211, 406–414.
Pickert, M. (1988). In vitro germination and storage of trinucleate
Arabidopsis thaliana (L.) pollen grains. Arabidopsis Inf. Serv. 26,
39–42.
Praefcke, G.J.K., and McMahon, H.T. (2004). The dynamin superfamily: Universal membrane tubulation and fission molecules? Nat. Rev.
Mol. Cell Biol. 5, 133–147.
Samuels, A.L., Giddings, T.H., Jr., and Staehelin, L.A. (1995). Cytokinesis in tobacco BY-2 and root tip cells: A new model of cell plate
formation in higher plants. J. Cell Biol. 130, 1345–1357.
Sanderfoot, A.A., Pilgrim, M., Adam, L., and Raikhel, N.V. (2001).
Disruption of individual members of Arabidopsis syntaxin gene families indicates each has essential functions. Plant Cell 13, 659–666.
Sano, T., Higaki, T., Oda, Y., Hayashi, T., and Hasezawa, S. (2005).
Appearance of actin microfilament ‘twin peaks’ in mitosis and their
function in cell plate formation, as visualized in tobacco BY-2 cells
expressing GFP-fimbrin. Plant J. 44, 595–605.
Schrick, K., Mayer, U., Horrichs, A., Kuhnt, C., Bellini, C., Dangl, J.,
Schmidt, J., and Jürgens, G. (2000). FACKEL is a sterol C-14
reductase required for organized cell division and expansion in
Arabidopsis embryogenesis. Genes Dev. 14, 1471–1484.
Torres-Ruiz, R.A., and Jürgens, G. (1994). Mutations in the FASS gene
uncouple pattern formation and morphogenesis in Arabidopsis development. Development 120, 2967–2978.
3518
The Plant Cell
Traas, J., Bellini, C., Nacry, P., Kronenberger, J., Bouchez, D., and
Caboche, M. (1995). Normal differentiation patterns in plants lacking
microtubular preprophase bands. Nature 375, 676–677.
Twell, D., Yamaguchi, J., and McCormick, S. (1990). Pollen-specific
gene expression in transgenic plants: Coordinate regulation of two
different tomato gene promoters during microsporogenesis. Development 109, 705–713.
Ursin, V.M., Yamaguchi, J., and McCormick, S. (1989). Gametophytic
and sporophytic expression of anther-specific genes in developing
tomato anthers. Plant Cell 1, 727–736.
Van Aelst, A.C., Pierson, E.S., Van Went, J.L., and Cresti, M. (1993).
Ultrastructural changes of Arabidopsis thaliana pollen during final
maturation and rehydration. Zygote 1, 173–179.
Van Damme, D., Bouget, F.-Y., Van Poucke, K., Inzé, D., and
Geelen, D. (2004a). Molecular dissection of plant cytokinesis and
phragmoplast structure: A survey of GFP-tagged proteins. Plant J. 40,
386–398.
Van Damme, D., Van Poucke, K., Boutant, E., Ritzenthaler, C., Inzé,
D., and Geelen, D. (2004b). In vivo dynamics and differential microtubule-binding activities of MAP65 proteins. Plant Physiol. 136, 3956–
3967.
Vanstraelen, M., Van Damme, D., De Rycke, R., Mylle, E., Inzé, D.,
and Geelen, D. (2006). Cell cycle dependent targeting of a kinesin at
the plasma membrane demarcates the division site in plant cells. Curr.
Biol. 16, 308–314.
Völker, A., Stierhof, Y.-D., and Jürgens, G. (2001). Cell cycle-independent expression of the Arabidopsis cytokinesis-specific syntaxin
KNOLLE results in mistargeting to the plasma membrane and is not
sufficient for cytokinesis. J. Cell Sci. 114, 3001–3012.
Wick, S.M. (1991). Spatial aspects of cytokinesis in plant cells. Curr.
Opin. Cell Biol. 3, 253–260.
Williamson, R.E., Burn, J.E., Birch, R., Baskin, T.I., Arioli, T., Betzner,
A.S., and Cork, A. (2001). Morphology of rsw1, a cellulose-deficient
mutant of Arabidopsis thaliana. Protoplasma 215, 116–127.
Yoneda, A., Akatsuka, M., Kumagai, F., and Hasezawa, S. (2004).
Disruption of actin microfilaments causes cortical microtubule disorganization and extra-phragmoplast formation at M/G1 interface in
synchronized tobacco cells. Plant Cell Physiol. 45, 761–769.
Zuo, J., Niu, Q.-W., Nishizawa, N., Wu, Y., Kost, B., and Chua, N.-H.
(2000). KORRIGAN, an Arabidopsis endo-1,4-b-glucanase, localizes
to the cell plate by polarized targeting and is essential for cytokinesis.
Plant Cell 12, 1137–1152.
Somatic Cytokinesis and Pollen Maturation in Arabidopsis Depend on TPLATE, Which Has
Domains Similar to Coat Proteins
Daniël Van Damme, Silvie Coutuer, Riet De Rycke, Francois-Yves Bouget, Dirk Inzé and Danny Geelen
Plant Cell 2006;18;3502-3518; originally published online December 22, 2006;
DOI 10.1105/tpc.106.040923
This information is current as of December 18, 2012
Supplemental Data
http://www.plantcell.org/content/suppl/2006/12/15/tpc.106.040923.DC1.html
References
This article cites 75 articles, 33 of which can be accessed free at:
http://www.plantcell.org/content/18/12/3502.full.html#ref-list-1
Permissions
https://www.copyright.com/ccc/openurl.do?sid=pd_hw1532298X&issn=1532298X&WT.mc_id=pd_hw1532298X
eTOCs
Sign up for eTOCs at:
http://www.plantcell.org/cgi/alerts/ctmain
CiteTrack Alerts
Sign up for CiteTrack Alerts at:
http://www.plantcell.org/cgi/alerts/ctmain
Subscription Information
Subscription Information for The Plant Cell and Plant Physiology is available at:
http://www.aspb.org/publications/subscriptions.cfm
© American Society of Plant Biologists
ADVANCING THE SCIENCE OF PLANT BIOLOGY